Mass spectrometry (MS) is an excellent technique for quantifying the amount of known and unknown substances in complex mixtures. Although, MS does have issues that can adversely affect the ability to assay or measure chemical and biological analytes. One problematic issue is ion suppression or enhancement in electrospray ionization (ESI), atmospheric pressure chemical ionization (APCI), and matrix assisted laser desorption ionization (MALDI) that are caused by nonvolatile compounds including salts, ion-pairing agents, endogenous compounds, and co-eluting compounds present in samples being evaluated by mass spectrometry. These ion suppressing or enhancing factors, referred to as sample matrix, cause changes in the efficiency of droplet formation or evaporation during the ionization process and ultimately result in an altered amount of charged analytes that reach the detector of the mass spectrometer. To illustrate this point, T. M. Annesley showed a signal response curve for caffeine in the absence and presence of serum extracts fractionated by high-pressure liquid chromatography (HPLC) coupled online to a mass spectrometer that showed a reduction in the signal for caffeine in the serum samples of 90% compared to caffeine reference solutions of equal concentration (Annesley, T M, et al Clin Chem, 2003; 49: 1041-1044). To address ion suppression or enhancement in liquid chromatography coupled to mass spectrometric assays internal standards have been used including stable isotopes of the compounds of interest or a compound that co-elutes with the compound of interest whereby the ion suppression or enhancement for both compounds is identical (Kitamura, R, et al J Chromatogra B Biomed Sci Appl, 2001; 754: 113-119), in the case of using a co-eluting compound to quantify the compound of interest, a single compound is used as the standard based on experimental knowledge of the retention characteristics.
Another method commonly used to account for matrix effects is the standard addition method (SAM), (Saxberg, B, et al Anal Chem, 1979; 51: 1031-1038). The SAM method works by measuring the analyte of interest in a sample with and without spiking in known amounts of the analyte. This generates a standard curve which can be extrapolated to determine the original analyte concentration. Despite SAM's widespread use for targeted measurements of specific compounds, it cannot be easily applied to proteomics or high-throughput experiments because investigators often lack a priori knowledge of what they wish to measure, purchasing standards for all the peptides of interest is prohibitively expensive, or the requirement of doing multiple measurements using the SAM is too costly.
AQUA peptides are isotopically labeled protein or peptide standards made specifically to quantitate a targeted protein or proteins of interest (Keshishian, H, et al Mol Cell Proteomics, 2007; 6: 2212-2229). AQUA peptides contain an isotopic mass altering atom(s) within a peptide that is diagnostic for a specific targeted protein. The AQUA peptide behaves chemically identical to the unlabeled peptide in biological samples and elutes chromatographically from reverse phase chromatographic separations at an identical retention time as the peptide from the biological sample. The AQUA peptide and the biological peptide can be differentiated by the mass spectrometer due to their mass difference. The concentration of AQUA peptide or protein is known and allows for the determination of the levels of proteins or peptides in a sample. The drawbacks to AQUA peptides include the necessity of having to purchase a separate AQUA peptide(s) for each protein to be measured from a biological sample, in some cases without experimental knowledge of the specific peptides mass spectrometry observability including limit of detection and limit of quantitation values.
A proteomics methodology referred to as QconCAT, or QCAT proteins, consist of concatenated tryptic peptides present in proteins that are being quantitatively measured in biological samples (Beynon, R, et al Nat Methods, 2005; 2; 587-589; Pratt, J M, et al Nat Protoc, 2006; 1: 1029-1043). QCAT proteins are designed to encode a set of peptides sequences that when subjected to endoproteinase cleavage are diagnostic for measuring the abundance of a targeted set of proteins in experimental samples. The gene encoding QCAT proteins are inserted into an expression vector and transformed into coli in minimal media containing 15NH4Cl as the nitrogen source. The amounts of target proteins in samples are determined by introducing a known amount of the isotopically labeled QCAT protein into the biological samples and co-digesting with an appropriate endopeptidase and comparing the ratios of the unlabeled biological peptides to the isotopically labeled QCAT peptides by mass spectrometry. Two major shortcomings of this approach are the need to develop a QCAT gene for each set of proteins to be quantified, and the potential for differential proteolysis occurring to the non-native form of the QCAT peptides as compared to the native peptides within their respective natural proteins.
QCAL1 artificial proteins (quantitative calibration artificial proteins) have also been used thr calibrating and defining instrument conditions (Eyers, C E, et al J Am Soc Mass Spectrom, 2008; 19: 1275-1280). The QCAL1 artificial protein was constructed using the same methodology as the QCAT proteins and consists of 22 tryptic peptide sequences designed to assess and optimize mass spectrometry resolution, mass calibration, linearity of signal detection, peptide separation by chromatography, and alignment of chromatograms from data collected over time from many LC/MS runs. These artificial proteins differ from the proposed technology because of their limited elution range, which results in these proteins not being generally applicable to correct for matrix effects. More specifically, application of the QCAL1 peptides involves their use to normalize for sample analytes that share no amount of co-fractionation with the QCAL1 peptides. The main drawback to using the QCAL1 peptides is that the amount of ion suppression or enhancement can vary dramatically throughout an LC/MS run (Stahnke, H, et al J Anal Chem, 2009; 81: 2185-2192).
Halogenated peptides as internal standards (H-PINS) have been used when mass spectrometry analysis is preceded by liquid chromatography (Mirzaei, H, et al J Mol Cell Prot, 2009; 8: 1934-1946). These internal standards contain halogen atoms that have V-shaped MS1 spectra that can be used to distinguish these internal standards from peptides present in complex matrices. A group of 10 H-PINS with a broad elution range spanning an LC/MS run has been used to perform instrument calibration, estimate overall sample matrix effects, and construct chromatographic alignment maps. The shortcoming of using a limited number of peptide standards over a broad elution range to normalize for ion suppression or enhancement is that ion suppression can change rapidly during an LC separation (Stahnke, H, et al J Anal Chem, 2009; 81: 2185-2192).
Another method used to correct for mass spectrometry detection efficiency is postcolumn infusion of standards (Stahnke, H, et al J Anal Chem, 2009; 81: 2185-2192). In this procedure, a single standard is continuously added to the effluent of an LC column immediately prior to the MS ionization source. This technique permits the assessment of analyte signal suppression or enhancement by different co-eluting matrix components during an entire chromatographic separation. Infusion of standards with very different chemical properties has been found to yield similar matrix effect responses, suggesting that such surrogate standards accurately report sample matrices. Postcolumn infusion has the drawback of requiring additional pumps or gradient formers and reduced sensitivity caused by dilution of the sample with the postcolumn infusate.
Another method used to study differences in the proteomes of cells in culture is referred to as Stable Isotope Labeled Amino acids in Cell culture (SILAC) (Ong, S E, et al Mol Cell Proteomics, 2002; 1: 376-386). SILAC works by metabolic incorporation of an isotopically modified light or heavy amino acid into proteins. In SILAC experiments, two or more groups of cells are grown in separate culture media with different light and heavy isotopic forms of a particular amino acid. One of the cell groups may be a control and the others represent a disease state or treated groups of cells. The cell lysates from the controls and disease states are mixed and proteolytically digested to produce peptides that behave chemically identical during liquid chromatography fractionation resulting in their co-elution. The relative abundance of proteins can be determined from the relative peak intensities observed in the MS or MS/MS spectrum for different cell cultures.
Despite the utility of SILAC there exist several shortcomings. The interpretation of data is difficult because each proteolytic peptide containing heavy or light amino acids generated from different samples are combined prior to mass spectrometric analysis resulting in different molecular ions and fragment ions depending upon the number of heavy or light amino acids in each peptide (Mellwain, S, et al Bioinformatics, 2008; 24; 339-347). Additionally, since different isotopes are being used it is necessary to reverse label control and disease samples to ensure that the isotopes themselves do not cause protein abundance changes. For instance, the metabolic conversion of arginine to proline in eukaryotes has been documented in SILAC experiments which causes incorrect quantitation by MS (Hwang, S I, et al Mol Cell Proteomics, 2006; 5; 1131-1145) and because more than 50% of tryptic peptides in large data sets contain proline (Van Hoof, D, et al Nat Methods, 2007; 4: 677-678) this is a significant problem for quantitative proteomic measurements. The final drawback for SILAC is that the methodology cannot be used to study tissue samples from animal and human sources.
Isobaric tags for relative and absolute quantitation (iTRAQ) is an amine-specific peptide based labeling method. iTRAQ has been used to study proteomes by data-dependent (Ross, P L, et al Mol Cell Proteomics, 2004; 3: 1154-1169) and MRM mass spectrometry (Wolf-Yadlin, A, et al Proc Natl Acad Sci USA, 2007; 104: 5860-5865). There are 3 different functional areas of each iTRAQ reagent including a reactive group, balancing group, and reporter group. The iTRAQ reagents are available with 8 different reporter peptide masses whose mass differences are offset by the balancing group, so that they all have the same overall mass. Thus, up to 8 different samples can be mixed together, each tagged with a different iTRAQ reagent. Because each tag has the same overall mass, peptides common to each biological sample will be detected at the same m/z; however, on subsequent fragmentation by MS/MS, the reporter region of each iTRAQ tag is separated from the balancing group which allows determination of the sample origin abundance of each fragmented iTRAQ peptide. The relative peak intensities of the reporter mass variants of a peptide reflects the relative concentration of a particular peptide in different samples.
The drawbacks with iTRAQ include variability among samples caused by the extra steps involved in labeling with the iTRAQ reagent. Also, the sensitivity and quantitative capability with iTRAQ is greatly reduced due to the combining of samples and corresponding increase in sample complexity. This increase in sample complexity often necessitates analyzing samples multiple times by mass spectrometry and generating an exclusion list of the most abundant ions. Peptides in this list are ignored for MS/MS analysis, so that less abundant peptides are more readily sequenced. In addition, since the iTRAQ reagent reacts with primary amines, peptides not containing lysine will be labeled at the N-terminus only, whereas lysine containing peptides will be labeled at both the N-terminus and on the lysine. If the labeling reactions are incomplete, multiple products will be generated which complicate the quantitative analysis. A statistical evaluation of these iTRAQ labeling concerns has revealed that at least a 2 fold change in peptide iTRAQ ratios is required to be biologically relevant (Armenta, J M, et al J Am Soc Mass Spectrom, 2009; 20: 1287-1302). In addition, iTRAQ is limited in utility to mass spectrometric instruments which have the ability to measure a specific precursor ion followed by fragmentation of the precursor ion commonly referred to as MS/MS.
The ICAT reagent also isotopically tags proteins, in this case by reacting with cysteine residues (Gygi, S P, et al Nat Biotechnol, 1999; 17: 994-999). Samples to be compared are individually reduced, denatured, and labeled potentially providing a source of differential error prior to mixing the different samples. The ICAT labeled samples are then subjected to 1D SDS-PAGE, in-gel trypsin digestion, peptide extraction, and purification of the tagged peptides through binding of the biotin moiety incorporated into the tag to avidin. Following cleavage of the biotinavidin tag, the peptides are analyzed by LC/MS/MS. In addition to the possible variability introduced by the steps involved in labeling each individual sample, ICAT has the drawback of labeling only cysteine containing peptides which limits analysis to only a small fraction of sample peptides.
Chromatographic separation with reverse phase liquid chromatography of peptides coupled to mass spectrometry has been used in many high-throughput proteomic and targeted protein analyses. Reverse phase liquid chromatography is utilized prior to mass spectrometry to partially fractionate complex peptide mixtures, thereby increasing the number of peptides that can be identified during an LC/MS/MS experiment (de Godoy, et al Genome Biology, 2006; 7: R50).
In non-isotopically labeled LC-MS and LC-MS/MS proteome experiments the method of spectral counting is frequently used to obtain correlative information on quantitative differences among proteins in different samples (Washburn, M P, et al Nat Biotechnol, 2001; 19: 242-247). Spectral counting refers to the number of times that a particular peptide is selected for fragmentation MS/MS by the mass spectrometer, which in turn is a function of its abundance relative to other co-eluting peptides. An alternative to spectral counting in label free proteomics is comparison among different samples of the area under the MS signal peak of a particular peptide (Bondarenko, P V, et al Anal Chem, 2002; 74: 4741-4749). A major advantage of the label free comparative proteomic methods is that in contrast to isotope labeling methods, there are no addition steps in the sample preparation process and samples are not overwhelmed by high abundance proteins preventing the detection and quantitation of low abundance peptides by being combined prior to MS analysis. A major drawback for the label free methods are that there is no compensation for differences in the suppression or enhancement of peptides caused by the variable makeup of different sample matrix components.